The yin and yang of hepatitis C: synthesis and decay of hepatitis C virus RNA.

Nature Reviews Microbiology | AOP, published online 10 August 2015; doi:10.1038/nrmicro3506

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The yin and yang of hepatitis C: synthesis and decay of hepatitis C virus RNA You Li1*, Daisuke Yamane1*, Takahiro Masaki1,2 and Stanley M. Lemon1,3

Abstract | Hepatitis C virus (HCV) is an unusual RNA virus that has a striking capacity to persist for the remaining life of the host in the majority of infected individuals. In order to persist, HCV must balance viral RNA synthesis and decay in infected cells. In this Review, we focus on interactions between the positive-sense RNA genome of HCV and the host RNA-binding proteins and microRNAs, and describe how these interactions influence the competing processes of viral RNA synthesis and decay to achieve stable, long-term persistence of the viral genome. Furthermore, we discuss how these processes affect hepatitis C pathogenesis and therapeutic strategies against HCV.

Lineberger Comprehensive Cancer Center, Division of Infectious Diseases, Department of Medicine and Department of Microbiology & Immunology, The University of North Carolina at Chapel Hill, Chapel Hill, North Carolina 27517–7292, USA. 2 Present address: Department of Virology II, National Institute of Infectious Diseases, 1‑23‑1 Toyama, Shinjuku‑ku, Tokyo 162–8640, Japan. 3 8.034 Burnett-Womack CB #7292, The University of North Carolina at Chapel Hill, Chapel Hill, North Carolina 27599–7292, USA. *These authors contributed equally to this work. Correspondence to S.M.L. e-mail: [email protected] doi:10.1038/nrmicro3506 Published online 10 August 2015 1

The pathogenesis of hepatitis C is intimately linked to the capacity of hepatitis C virus (HCV) to persist for decades in most infected persons. The mechanisms underlying this long-term HCV persistence remain incompletely characterized but include multiple viral assaults on the host innate immune system coupled with defective adaptive immunity 1,2. Although interferon (IFN)-based therapies, which are at best only modestly effective, are being rapidly supplanted by highly potent IFN-sparing regimens containing multiple direct-acting antiviral agents (DAAs)3 (BOX 1), HCV will remain a public health concern for many years. Many individuals are unaware of their infection, and access to curative therapies is limited by a variety of economic and social issues4. In addition, those fortunate enough to be cured of chronic hepatitis C with DAAs may remain at risk for liver cancer for an indefinite period. Approximately one in three acutely infected persons spontaneously clears HCV infection, but in most individuals, HCV persists within the liver for years with minimal pathology and relatively constant levels of viraemia5. Spontaneous clearance is closely associated with a polymorphism in the IFNL4 gene (which encodes IFNλ4)6. For reasons that are poorly understood but that may also involve cell-intrinsic antiviral defences, infection is restricted to a minority of hepatocytes with a patchy distribution in the liver 7–9. The intracellular abundance of viral RNA and proteins is low, making the identification of these cells difficult. Pathology, exemplified primarily by increasing intrahepatic fibrosis,

develops insidiously and is caused largely, if not entirely, by immune responses to the virus10,11. The ability of the virus to restrict infection to a minority of cells and to maintain a low intracellular abundance of viral RNA and proteins serves the virus well but demands close regulation of the replicative machinery. For example, overly abundant production of new viral genomes and viral proteins could lead to apoptotic cell death, as observed in cell cultures infected with laboratory strains of HCV12,13. By contrast, less efficient replication might allow the virus to be eradicated by a combination of innate and adaptive immune responses. Long-term persistence thus requires a balance between the synthesis and decay of viral RNA in infected cells. In part, this balance is achieved by a unique capacity of the HCV replicase to be negatively regulated by virus-induced oxidative membrane damage14. Whereas infection induces oxidative stress, the ability of HCV to replicate is restricted by the peroxidation of lipids associated with its replication machinery. This may contribute to persistence by maintaining viral RNA and antigen synthesis at low levels, and preventing virusinduced cytopathology. Adding to this balancing act is an unusual adaptation: the viral genome is stabilized and its degradation prevented by interactions with an abundant, liver-specific microRNA (miRNA), miR‑122 (also known as hsa-miR‑122‑5p)15. In this Review, we describe the factors that control the balance between HCV RNA synthesis and decay and discuss how this balance affects both viral pathogenesis and therapeutic strategies aimed at eliminating HCV infection.

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REVIEWS Box 1 | Host-targeting and direct-acting antivirals miR-122 antagomir

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Reviews | Microbiology Unlike human immunodeficiency virus 1 (HIV‑1), hepatitis C virus (HCV) does not integrate Nature its genome within the host genome, and infections can therefore be completely cured and the virus eradicated by effective antiviral therapies. Before 2011, the standard of care for the treatment of hepatitis C was a combination of weekly injections of pegylated recombinant interferon‑α and daily oral ribavirin. This therapy was continued for periods of 6–12 months and was often associated with substantial side effects. Cure (a sustained virological response (SVR)) was achieved in at most 50% of patients infected with the most difficult-to-treat HCV genotype, genotype 1. However, the outlook for infected patients has been radically altered in the past 5 years with the introduction of direct-acting antivirals (DAAs; see the figure; brown boxes). These small molecules target specific proteins expressed by HCV, can be administered orally and (with newer formulations) have minimal side effects. There are four major classes of DAAs3. Two of these classes target active sites in viral enzymes that are essential for replication: the NS3‑4A protease and the NS5B polymerase. A third major class targets the NS5A protein, inhibiting NS5A activities in both biogenesis of the membranous web and virus assembly38,39. The fourth class comprises allosteric inhibitors that target several sites in the NS5B polymerase. Drugs in each class have now been approved for use by the US Food and Drug Administration. An overarching problem for DAAs is their ability to rapidly select for resistant viruses, which is due to the highly replicative nature of HCV infection148 and the error-prone RNA synthesis that typifies positive-strand RNA viruses. However, this problem has been largely overcome by multidrug regimens that combine two or more DAAs. Thus, currently available all-oral combination therapies taken for 12 weeks (and possibly less time) provide SVR rates of ~95% for patients with HCV genotype 1 infections. Although the currently available DAAs are limited in their coverage of other genotypes — particularly HCV genotype 3, which is increasingly common in Europe — second-generation DAAs that are now in development may ameliorate this issue. Host-targeting antivirals represent an alternative approach, to avoid resistance and achieve pan-genotypic coverage (see the figure; green boxes). This strategy is best exemplified by non-immunosuppressive inhibitors of cellular cyclophilins52, and by an injected oligonucleotide (an antagomir) that binds to and sequesters the microRNA miR‑122 (REFS 85,96). Both cyclophilins and miR‑122 are important host factors that participate in HCV replication. These host-targeting strategies have shown promise in clinical trials, but neither has achieved regulatory approval.

Stem–loop Also called a hairpin element. An element of RNA secondary structure formed when two segments of an RNA molecule have complementary nucleotide sequences and base pair to form a two-stranded helix with an intervening single-stranded loop segment.

Internal ribosome entry site (IRES). A highly structured RNA sequence that recruits ribosomes and initiates translation of the RNA independently of ribosome scanning from a 5ʹ‑cap. The 5ʹ untranslated region of the hepatitis C virus RNA genome contains an IRES. A variety of IRES elements with different RNA structures are found in other viruses, as well as in a subset of host mRNAs, where they permit translation when cap-dependent translation is inhibited during mitosis, apoptosis or hypoxia.

The yin of hepatitis C: viral RNA synthesis The HCV life cycle has been studied intensively, and how the virus enters cells, assembles into new virions and is released from cells has been reviewed elsewhere16,17. Here, we focus on the organization of the viral genome and discuss the viral and cellular factors that are involved in HCV RNA synthesis. HCV genome organization. HCV is a positive-strand RNA ((+)RNA) virus with a 9.7 kb single-stranded, messenger-sense RNA genome that is translated directly following its release into the cytoplasm. The RNA contains a lengthy open reading frame (ORF) flanked by highly structured terminal untranslated regions (UTRs) that are essential for viral RNA synthesis and translation18–22 (FIG. 1a). Unlike canonical cellular mRNAs, the viral genome lacks a 5ʹ terminal 7‑methylguanylate cap (m7GpppN), terminating instead in a 5ʹ triphosphate23. It also lacks a 3ʹ poly(A) tail, terminating in a conserved, stable stem–loop structure21. The ORF is translated under the control of a type III internal ribosome entry site (IRES) located within the 5ʹ UTR24. Translation results in the synthesis of a large polyprotein that is processed by cellular and viral proteases to yield three structural

proteins (core, envelope 1 (E1) and E2), which are components of the virion, and seven nonstructural proteins (p7, NS2, NS3, NS4A, NS4B, NS5A and NS5B), some of which have distinct enzymatic activities25 (FIG. 1b). Selfreplicating RNA replicons require only NS3, NS4A, NS4B, NS5A and NS5B for efficient viral RNA replication26. These nonstructural proteins assemble as a replicase complex in association with vesicle-like membranes called the ‘membranous web’, which is the presumed site of viral RNA synthesis27,28 (FIG. 2). Because viral RNA synthesis is directed by the membrane-bound replicase complex, it requires prior translation of the viral genome and expression of the nonstructural proteins. Genome amplification commences with primer- independent initiation of the synthesis of a negativestrand RNA ((–)RNA); complement of the (+)RNA) genome, catalysed by NS5B, an RNA-dependent RNA polymerase. Recent crystallographic studies of the polymerase stalled at various stages of initiation provide a dynamic view of this process29. A β‑loop and a carboxy‑terminal membrane-anchoring linker restrict access to the polymerase active site in the apo state but retract to accommodate and position the 3ʹ end of the (+)RNA template, together with incoming nucleotides,

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Figure 1 | The hepatitis C virus RNA genome and polyprotein. a | Secondary RNA structure within the single-stranded Nature Reviews | Microbiology 9.7 kb positive-strand RNA ((+)RNA) genome of hepatitis C virus (HCV), showing the short, highly structured 5ʹ and 3ʹ untranslated regions (UTRs; black) that flank the ends of the internal long open reading frame (ORF) encoding the viral polyprotein (blue and orange, see part b). The 5ʹ UTR contains essential RNA replication signals, including the two binding sites (S1 and S2, red arrows) for the microRNA miR‑122 and an internal ribosome entry site (IRES) that contains multiple pseudoknots (PKs) and binds 40S ribosome subunits with high affinity. The 3ʹ UTR contains a highly variable poly(U/UC) tract followed by the nearly absolutely conserved 98 nucleotides that constitute the 3ʹ X‑tail sequence at the 3ʹ terminus; both of these sequences are essential for RNA replication. Initiator AUG and terminator UGA codons at the ends of the ORF delimit non-coding and coding RNA sequences. Also shown is the stem–loop (SL) 5BSL3.2, the core of a conserved compound RNA structure; this structure is located near the 3ʹ end of the ORF and functions as a cis-acting replication element (CRE). Additional, less conserved secondary RNA structure is also present within the polyprotein-coding region. The coloured lines represent a network of dynamic long-range RNA–RNA interactions both within and between highly structured RNA domains at the ends of the genome. SLs other than the CRE are labelled according their position within the genome. b | The ORF encodes a single large polyprotein that is processed by cellular and viral proteases into ten mature viral proteins: three that are structural and present in the virion, and seven nonstructural proteins. Amino acid numbers in the polyprotein are indicated. The structural proteins (blue) are the viral core protein and two glycosylated envelope proteins, E1 and E2, and are derived from the amino terminus of the polyprotein. Nonstructural proteins required for RNA synthesis (orange) include NS3 (a protease and helicase), NS4A (a protease cofactor), NS4B (an integral membrane protein), NS5A (involved in replicase formation and virion assembly) and NS5B (an RNA-dependent RNA polymerase (RdRp)); regions of these proteins with specific enzymatic activities are indicated below the polyprotein schematic. The nonstructural proteins p7 (an ion channel) and NS2 (a cis-acting protease and assembly factor; both shown in green) are essential for virion morphogenesis and egress, but not for RNA replication. All of these proteins are likely to be multifunctional. Arrowheads indicate sites of cleavage by the viral proteases NS2‑3 (purple) and NS3‑4A (red). Part a is adapted from Romero-Lopez, C. et al. End to end crosstalk within the hepatitis C virus genome mediates the conformational switch of the 3’ X tail region. Nucleic Acids Res. 2014, 42, 567–582, by permission of Oxford University Press.

within the active site. This ensures that (–)RNA synthesis begins at the extreme 3ʹ terminus of the genome. Following the slow formation of a dinucleotide, which acts as primer, additional conformational changes in NS5B allow egress of the elongating RNA chain, with the polymerase proceeding in a 3ʹ‑to‑5ʹ direction along the length of the (+)RNA genome29 (FIG. 2). As

translating ribosomes move in the opposite direction (that is, in the 5ʹ‑to‑3ʹ direction), they present an insurmountable barrier to the replicase, and translation must first be terminated before an RNA molecule can serve as a template for RNA synthesis. The factors regulating this switch from RNA translation to RNA synthesis are poorly understood.



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Lipid droplets Ubiquitous, dynamic cellular organelles that are rich in lipids and mainly composed of cholesteryl esters and triglycerides. They function in energy storage by sequestering fatty acids in triglycerides and as a platform for the metabolism and transport of lipids, as well as playing a part in cellular signalling through the generation of bioactive lipids.

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Figure 2 | The yin of hepatitis C virus: RNA synthesis. On hepatitis C virus (HCV) entry into theReviews infected|cell, the viral Nature Microbiology positive-strand RNA ((+)RNA; red) is released into the cytoplasm. (+)RNA from entering virions must first interact with ribosomes in the endoplasmic reticulum (ER) to undergo translation in order to produce the viral nonstructural proteins that are required for the assembly of replicase complexes. These complexes are associated with membranes derived from the ER and, together with the membranes, form the membranous web. Within the membranous web, (+)RNA is copied to a complementary negative-strand RNA ((–)RNA) intermediate (blue), which serves as the template for the synthesis of multiple new (+)RNA strands. These progeny RNA molecules have four possible fates: they may cycle to rough ER for translation and the production of additional viral proteins; they may be shuttled to lipid droplets (LDs) for packaging into nascent virions; they may be retained within the membranous web to provide a template for the synthesis of additional (–)RNA; or they may undergo intracellular decay (such as 5ʹ‑to‑3ʹ exoribonuclease 1 (XRN1)‑mediated decay). Factors regulating the movement of the viral RNA between compartments are poorly understood. The overall replicative cycle induces oxidative stress, with subsequent oxidative membrane damage that acts to restrict the efficiency of the membranous web (not shown). The inserts represent detailed schematics of viral RNA translation and synthesis. (+)RNA molecules can engage in either protein synthesis (left) or RNA synthesis (right), but not both simultaneously owing to the conflicting directions of movement for ribosomes (5ʹ to 3ʹ) and the replicase (3ʹ to 5ʹ) on (+)RNA. Figure is adapted with permission from REF. 98, Elsevier.

Newly synthesized (–)RNA molecules then serve as templates for the production of new (+)RNA molecules, which have several possible fates including: moving to the endoplasmic reticulum (ER) to serve as mRNA for

continuing viral protein synthesis; trafficking to lipid droplets, which serve as a platform in the assembly of progeny virions to be exported from the cell; remaining within the membranous web as templates for additional



REVIEWS (–)RNA synthesis (although it is possible that a (+)RNA molecule must first be translated to become competent as a template for (–)RNA synthesis); and undergoing degradation by host ribonucleases (FIG. 2).

Retinoic acid-inducible gene I (RIG‑I). A cytoplasmic RNA helicase that senses viral RNAs and initiates signalling that leads to host interferon production. RIG‑I preferentially binds short double-stranded, AU‑rich RNAs with 5ʹ dior triphosphates and subsequently activates its downstream adaptor, mitochondrial antiviral signalling protein (MAVS), through interactions between shared caspase activation and recruitment domains (CARDs). RIG‑I and a second RIG‑I‑like receptor, MDA5 (also known as IFIH1), contribute to innate immune sensing of distinct types of viruses.

Protein kinase R (PKR). An interferon-induced protein kinase that is activated in response to double-stranded RNA and is central to cellular responses to a variety of stresses, such as viral infection, cytokine exposure, nutrient depletion, irradiation and endoplasmic reticulum stress. PKR-mediated phosphorylation of eukaryotic translation initiation factor 2A (EIF2A) results in global translational repression.

Pseudoknot A higher-order RNA structure that contains at least two stem–loops and in which the bases in one half of one stem pair with those in the loop of a second stem–loop.

Ribosome scanning The process by which a 40S ribosome subunit with associated initiation factors moves in a 3ʹ direction along an mRNA molecule in search of a start codon (typically AUG) following its initial interactions with and binding to the 5ʹ terminal 7‑methylguanylate cap structure. This is an essential component of eukaryotic cap-dependent translation initiation. Assembly of the mature 80S ribosome and the initiation of polypeptide synthesis commence at the start codon.

Architecture and composition of the HCV replicase. Cryoelectron microscopy of the membranous web reveals both single-membrane vesicles and double-membrane vesicles (DMVs) formed by protrusions of the ER membrane into the cytosol30. The presence of double-stranded RNA and nonstructural proteins within DMVs suggests that these vesicles are the site of viral RNA synthesis30–32, although conclusive proof for this is lacking. In addition to providing a platform for RNA synthesis and maintaining high local concentrations of essential nonstructural proteins, the DMVs may also contribute to viral immune evasion. For example, DMVs may sequester replication intermediates from cytosolic innate immunity sensors of non-self RNAs, such as retinoic acid-inducible gene I (RIG‑I; also known as DDX58) and protein kinase R (PKR). DMVs are enriched in cholesterol and sphingo lipids33, and their formation is dependent on ongoing sphingolipid synthesis34. The membranes of DMVs are relatively resistant to detergents and protect the vesicle contents from host ribonucleases and proteases33. Notably, although ectopic expression of NS4B alone induces distinct membrane alterations, NS5A expression is required for the formation of DMVs30,35. Thus, both NS4B and NS5A are likely to be involved in the biogenesis of the membranous web. In addition to viral proteins, cellular factors are also involved in biogenesis of the membranous web. For example, phosphatidylinositol‑4‑phosphate catalysed by phosphatidylinositol 4‑kinase IIIα (PI4KIIIα; also known as PI4Kα) regulates the formation and integrity of DMVs through a mechanism that is yet to be defined, whereas the activity of the kinase is stimulated by NS5A36,37. Consistent with this, DAAs targeting NS5A block assembly of the membranous web but have little or no effect on ongoing RNA synthesis by previously assembled replicase complexes38–40 (BOX 1). Although X‑ray crystallography has provided model structures for the entire NS3 molecule (in complex with part of NS4A)41, for the NS5B polymerase42 and for the amino‑terminal domain of NS5A43,44, very little is known about how these key molecules associate with each other within the replicase. NS3 is anchored to membranes via its N‑terminal domain, as well as via interactions with its membrane-bound NS4A cofactor, and NS5B is anchored to membranes via its C‑terminal domain25. NS5A is similarly anchored to membranes, most likely near a dimerization domain close to its N terminus, whereas NS4B is an integral membrane protein. Each of these proteins is required for viral RNA synthesis, and several play multiple parts in this process. For example, DAAs targeting the NS3‑4A protease not only block processing of the polyprotein (a process that is required for replicase assembly) but also directly inhibit RNA synthesis and virus assembly 39, suggesting a role for NS3 in each of these processes (BOX 1). Therefore, the nonstructural

proteins are multifunctional. Moreover, these proteins are expressed more abundantly than is required for the formation of the membranous web45. Substantial amounts of these proteins are present outside the membranous web, where they may engage in functions other than RNA synthesis. In addition to the role of PI4KIIIα in the biogenesis of the membranous web, other host proteins are required for HCV genome amplification. Peptidyl-prolyl cis–trans isomerase A (PPIA; also known as cyclophilin A), VAMP-associated protein A (VAPA) and VAPB contribute directly to RNA synthesis and are associated with the membranous web46,47. PPIA, an abundant protein, binds to NS5A and may facilitate the recruitment of NS5B into the replicase complex by modifying proline side chains near the membrane anchor at the NS5B C terminus48,49. VAPA and VAPB are vesicle-associated membrane proteins that have roles in sphingolipid metabolism and vesicle transport; they form homodimers and heterodimers and are required for assembly of the replicase complex 50,51. Like PPIA, VAPA and VAPB bind to NS5A and may also facilitate the incorporation of NS5B into the membranous web. Importantly, cells that are depleted of these proteins do not support HCV replication, and cyclophilin inhibitors have shown therapeutic promise against hepatitis C in humans52. Structure of HCV RNA. Highly structured but variably conserved RNA elements throughout the HCV genome contribute to the efficiency of RNA synthesis53 (FIG. 1). Although many of these RNA structures are incompletely characterized, it is thought that they may interact with both viral and host proteins, including components of the translation machinery and replicase complex. The structure of the 5ʹ UTR is well established. It is highly conserved and contains a small stem–loop near the 5ʹ terminus that is essential for RNA replication, possibly because it recruits a viral or host cell protein18,19. Larger, more complex stem–loops and a pseudoknot between nucleotides 45 and 355 form the IRES18,54, which binds the 40S ribosome subunit with high affinity in association with eukaryotic initiation factor 3 (eIF3) (FIG. 1). Translation initiates internally on the RNA without ribosome scanning and thus differs from the typical process in mammalian cells and superficially resembles events in prokaryotes. The 3ʹ UTR, including a highly conserved 3ʹ ‘X‑tail’ (the 3ʹ terminal 98 nucleotides) is also highly structured55 (FIG. 1). Also highly conserved, it contains several stem–loops and a U at the 3ʹ terminal position that are important for replication, as is an upstream poly(U/UC) tract of variable length and sequence20,56 (FIG. 1). Both the sequence and structure within these RNA elements are essential, but their exact roles in RNA replication are not known. Recent studies using selective 2ʹ‑hydroxyl acylation and primer extension (SHAPE) have revealed additional, less conserved RNA structures within internal regions of the genome53,57,58. These results extend earlier observations that support the functional importance of RNA structure within both the core and NS5B‑coding regions59,60. A stem–loop embedded within a cruciform structure



REVIEWS Selective 2ʹ‑hydroxyl acylation and primer extension (SHAPE). An experimental method that assesses the backbone flexibility of an RNA molecule at a single-nucleotide resolution based on the reactivity of 2ʹ‑hydroxyl groups towards an electrophile. Nucleotides that are flexible (unpaired) are preferentially chemically modified and are recognized as stops in a subsequent primer extension reaction, allowing unpaired bases to be distinguished from those for which flexibility is constrained by base-pairing.

RNAi A mechanism that was initially discovered in plants and involves small non-coding RNAs suppressing gene expression by targeting specific mRNA transcripts in association with an RNA-induced silencing complex. Synthetic short interfering RNAs (siRNAs) may be transfected into cells to deplete a specific protein.

in the NS5B‑coding region (called 5BSL3.2 or SL9266), which had been identified previously as an essential cis-acting RNA element (CRE)60, engages in mediumrange interactions with nucleotides located upstream, around nucleotide 9,033 (REF. 61) or nucleotide 9,110 (REF. 57), as well as engaging in a ‘kissing’ interaction with a stem–loop within the 3ʹ UTR62,63 (FIG. 1). Other RNA elements within the NS5B‑coding region and upstream of 5BSL3.2 also seem to be essential for efficient replication53,64. Longer-range interactions between the CRE, the 3ʹ UTR and the IRES have been identified and may influence the structure of both the IRES and the 3ʹ X-tail58,65. Thus, a network of variably conserved RNA–RNA interactions exists across the HCV RNA genome and between its 5ʹ and 3ʹ ends, and this may modulate various steps of the replication cycle. RNA–RNA interactions between the 5ʹ and 3ʹ ends may promote genome circularization, which is a common phenomenon among (+)RNA viruses and may either enhance translation, by facilitating the recycling of 40S ribosome subunits, or seed the initiation of (–)RNA synthesis (BOX 2). Host RNA-binding proteins. Numerous host RNAbinding proteins interact with the HCV genome and promote its replication. Those that bind the 5ʹ UTR include La protein (SSB), poly(rC)-binding protein 2 (PCBP2), polypyrimidine tract-binding protein 1 (PTBP1), PTBP2, heterogeneous nuclear ribonucleoprotein L (HNRNPL), insulin-like growth factor 2 mRNAbinding protein 1 (IGF2BP1; also known as IMP1), interleukin enhancer-binding factor 3 (ILF3; also known as NF90, NFAT90 or NFAR) and the U6 snRNA-associated Sm‑like (LSM) proteins LSM1–LSM7, whereas synaptotagmin-binding cytoplasmic RNA-interacting protein (SYNCRIP; also known as HNRNPQ or NSAP1) binds to an RNA sequence immediately downstream of

Box 2 | Circularization of viral positive-strand RNA genomes Genome circularization is a crucial step in the initiation of RNA synthesis for some positive-strand RNA viruses151 and can be mediated either by RNA–RNA interactions or RNA–protein–RNA interactions at the 5ʹ and 3ʹ ends of the genome. During replication of the ‘classical’ flaviviruses, including dengue virus and yellow fever virus, circularization is mediated by direct base pairing between complementary sequences near the 5ʹ and 3ʹ ends of the genome. Mutations in these regions impair RNA synthesis without affecting translation of the viral RNA. The 5ʹ end of the genome of these viruses contains a conserved Y‑shaped stem–loop structure (5ʹ SL) that is required for binding and activity of the RNA polymerase152. Thus, communication with the 5ʹ end is required for the initiation of negative-strand synthesis at the 3ʹ end. By contrast, circularization of the poliovirus genome is mediated by RNA–protein and protein–protein interactions between poly(A)-binding protein (PABP) bound to the 3ʹ poly(A) tail of the genome and both poly(rC)-binding protein 2 (PCBP2) and the viral 3CD protein bound to a 5ʹ cloverleaf-like structure in the viral RNA153. This enables the viral RNA-dependent RNA polymerase, 3Dpol, to position itself at the 3ʹ end of the genome and initiate the synthesis of negative-strand RNA. The long-range interaction with the ribonucleoprotein complex at the 5ʹ end of the genome may also induce conformational changes in the 3ʹ untranslated redion (3ʹ UTR) that are required for RNA synthesis. There is no conclusive proof for circularization of the HCV genome, but several lines of evidence suggest that this occurs and is mediated by PCBP2, interleukin enhancer-binding factor 3 (ILF3)70,72 and possibly other RNA-binding proteins that interact with both 5ʹ and 3ʹ UTRs (BOX 4).

the 5ʹ UTR66–71. In many cases, depletion of these proteins impairs HCV translation, suggesting they act as IRES-transactivating factors (ITAFs) and enhance the expression of the viral polyprotein. Several different molecular mechanisms are likely to underpin ITAF activity, and these may differ depending on the experimental system used. Most of these proteins bind both 5ʹ UTRs and 3ʹ UTRs, and as shown for PCBP2 (REF. 72), such proteins may promote circularization of the genome, thus favouring translation (BOX 2). Other possible roles of ITAFs include remodelling of ribonucleoprotein structure, leading to enhanced recruitment of the translation initiation factor eIF3, which binds a stem–loop within the IRES and is essential for HCV translation73. Other 5ʹ UTR-binding proteins, such as HNRNPL, may interact with RNA within the replication complex and thus directly influence RNA synthesis74. In most cases, a role for these proteins in protecting viral RNA against host nuclease-mediated degradation has not been studied, and this possibility cannot be excluded (see below). Additional host proteins bind the 3ʹ UTR. HNRNPC, ELAV-like 1 (ELAVL1; also known as HUR), glyceraldehyde‑3‑phosphate dehydrogenase (GAPDH) and either PTBP1 or PTBP2 bind to the poly(U/UC) tract 75–79. ELAVL1 and either PTBP1 or PTBP2 also bind the highly conserved 3ʹ X‑tail, and ELAVL1 also binds the 3ʹ end of the complementary HCV (–)RNA76. Although the functional importance of most of these specific protein–RNA interactions is not clear, experimental depletion of each of these proteins using RNAi impairs viral replication80. In the case of ELAVL1, this effect may be independent of the stabilizing actions that ELAVL1 possesses for cellular mRNAs76. In addition to proteins binding the 5ʹ and 3ʹ UTRs, Ewing sarcoma breakpoint region 1 (EWSR1; also known as EWS) promotes viral replication by binding to the CRE81. It does so preferentially in the absence of the kissing interaction between the CRE and the 3ʹ UTR, and it is recruited into the membranous web, which suggests that EWSR1 has a direct role in RNA replication. Interactions between the genome and host RNAbinding proteins may also affect HCV replication indirectly. For example, binding of the multifunctional ATP-dependent DEAD box RNA helicase DDX3X to the 3ʹ UTR activates conserved helix–loop–helix ubiquitous kinase (CHUK; also known as IKKα), which increases expression of sterol regulatory elementbinding proteins (SREBPs) through the CREB-binding protein (CBP; also known as CREBBP)–p300 pathway, thereby increasing the formation of the lipid droplets that serve as platforms for virus assembly 82. HCV infection alters the subcellular distribution of DDX3X such that it colocalizes with the viral core protein. As DDX3X plays a supporting part in RIG‑I‑mediated induction of IFNβ83, its interaction with the viral core could also modulate innate immune responses to HCV. Thus, the viral genome has evolved a set of sophisticated mechanisms to manipulate the cellular environment in multiple ways in order to facilitate productive replication.



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RNA-induced silencing complex (RISC). A ribonucleoprotein complex with a core comprising a small non-coding RNA (typically a microRNA or short interfering RNA) guide strand loaded into an Argonaute protein (AGO1– AGO4). The RISC acts to suppress translation and, in some cases, directly cleave an mRNA target.

Antagomir A synthetic, typically nucleaseresistant, chemically modified oligonucleotide that is complementary to the sequence of a microRNA and antagonizes its activity by binding to and sequestering the microRNA from its mRNA target, as well as possibly blocking microRNA biogenesis.

Small nucleolar RNA (snoRNA). A class of small non-coding RNAs that act to guide specific post- transcriptional modifications, such as 2ʹ‑O‑methylation or uridine isomerization, and occasionally cleavage, of other RNAs (primarily rRNAs and tRNAs).

miR‑122 and viral RNA synthesis. miR‑122 is an abundant liver-specific miRNA that regulates the expression of many genes, including those responsible for fatty acid and cholesterol metabolism84,85, and also has significant tumour suppressor activity 86. Whereas miRNAs typically regulate gene expression by binding to the 3ʹ UTR of mRNAs in association with an RNA-induced silencing complex (RISC), thereby downregulating translation and enhancing mRNA decay, miR‑122 binds to two tandem sites (S1 and S2) close to the 5ʹ end of the HCV genome and promotes its replication87,88 (FIG. 1). Binding of miR‑122 to S1 and S2 follows canonical miRNA binding patterns, with base-pair formation involving miRNA nucleotides 2–8 (the ‘seed sequence’) as well as downstream (that is, 3ʹ) bases89–91 (BOX 3). This is consistent with the binding of miR‑122 in association with Argonaute 2 (AGO2) as a RISC-like complex that may also contain trinucleotide repeat-containing 6A (TNRC6A) 15,92. High-throughput sequencing and crosslinking immunoprecipitation (HITS–CLIP) using an antibody against human AGO proteins has confirmed binding of miR‑122 to both S1 and S2 in the viral genome93. Other conserved potential seed sequencebinding sites, in the IRES and within the ORF, are also occupied by miR‑122, but the dominant miR‑122 interactions are at S1 and S2 (REFS 90,93). Although it is not the most abundant miRNA in hepatocytes, miR‑122 is the dominant miRNA binding to the viral genome. miR‑122 binding to both S1 and S2 is important for viral replication, and recent work supports simultaneous occupancy at both sites90,94,95. HCV RNAs with base substitutions within either S1 or S2 are impaired in replication but can be rescued by complementary miR‑122 mutants87,94. This indicates that miR‑122 promotes HCV replication not by modulating host gene expression but by direct interactions with the viral RNA. Highlighting the importance of miR‑122 in the HCV life cycle, an antisense locked nucleic acid antagomir called miravirsen, which binds and sequesters miR‑122, demonstrated substantial antiviral activity in preclinical and clinical trials85,96. Although some studies suggest that miR‑122 promotes HCV replication by stimulating translation94,97, more recent data support two distinct alternative mechanisms. First, miR‑122 protects HCV RNA against degradation mediated by 5ʹ‑to‑3ʹ exoribonuclease 1 (XRN1), a exoribonuclease involved in host mRNA decay 23 (see below). Equally (if not more) importantly, miR‑122 induces an increase in the rate of viral RNA synthesis when transfected into infected cells98. This increase in RNA synthesis can be measured by quantifying 5‑ethynyl uridine incorporated into nascent HCV RNA and is apparent within an hour of miR‑122 transfection. Increases in RNA synthesis precede any measurable increase in the incorporation of labelled amino acids into newly synthesized viral proteins, consistent with the primary effect of the miRNA being on the replicase98. The ability of miR‑122 to stimulate HCV RNA synthesis is dependent on AGO2 and is not observed in AGO2‑depleted cells98. This is further evidence that miR‑122 binds the RNA in association with AGO2 and is

consistent with other studies demonstrating the dependence of HCV replication on AGO2 (REFS 15,99,100). The miR‑122–AGO2 complex could also mask the presence of a 5ʹ triphosphate and thereby limit the recognition of HCV RNA by the immune sensor RIG-I101. However, this has not been experimentally demonstrated, and it is not likely to be the primary function of miR‑122, as miR‑122 promotes both HCV RNA stability and viral RNA synthesis in Huh‑7.5 cells that lack functional RIG‑I activity 15,98,102. Despite the importance of miR‑122 in the viral life cycle, some HCV mutants have reduced requirements for miR‑122. The S1 site is ablated in a novel virus in which the 5ʹ stem–loop is replaced with cellular U3 small nucleolar RNA (snoRNA) and, perhaps because miR‑122 continues to bind to S2, this virus replicates well in the absence of miR‑122 (REFS 93,103). HCV variants with substitutions at nucleotide 3 (C3U) or nucleotide 28 (G28A) are also relatively resistant to miR‑122 sequestration104,105. Low-level HCV replication has also been reported in cells lacking detectable miR‑122 (REF. 106). Whether specific compensatory mechanisms support HCV replication under these conditions is unknown. In addition to stabilizing HCV RNA and promoting its synthesis, the binding of miR‑122 to the viral RNA exerts a ‘sponge’ effect in cultured cells infected with robustly replicating laboratory strains of HCV, resulting in functional de‑repression of host mRNAs that are normally targeted by miR‑122 (REF. 93). A meta-analysis of published array data provides some evidence for this effect in HCV-infected cirrhotic liver tissue93, but it remains uncertain whether a biologically significant sponge effect occurs in vivo, where miR‑122 abundance is greater and HCV RNA abundance much lower than in infected cell cultures. Balancing HCV RNA translation versus synthesis. Given the conflict inherent in 5ʹ‑to‑3ʹ translocation of ribosomes on the HCV genome during protein synthesis versus the 3ʹ‑to‑5ʹ movement of the polymerase required for (–)RNA synthesis, a mechanism must exist for balancing the engagement of the genome in RNA translation versus RNA synthesis. Among other possibilities, miR‑122 seems to have a critical role in this process. The increases in viral RNA synthesis that are mediated by miR‑122 require active cellular protein synthesis and are blocked by treatment with cycloheximide or puromycin, both of which are potent general protein synthesis inhibitors98. However, puromycin alone enhances viral RNA synthesis in the short term, whereas cycloheximide alone does not 98. These results are best explained by the fact that puromycin induces the premature release of translating ribosomes from RNA, wheres cycloheximide does not. Thus, it is likely that puromycin stimulates RNA synthesis by freeing up viral genomes that are engaged in translation, making them available for viral RNA synthesis. Under these conditions, miR‑122 fails to further stimulate RNA synthesis, suggesting that miR‑122 acts similarly and thus redundantly to puromycin, stimulating viral RNA synthesis by increasing the fraction of genomes engaged in this process98. Consistent with this



REVIEWS Box 3 | Biogenesis of miR‑122 and its interactions with the hepatitis C virus genome MicroRNAs (miRNAs) are small duplex non-coding RNAs of ~22–23 nucleotides in length that have 2‑base 3ʹ overhangs. These small RNAs regulate the expression of eukaryotic genes by repressing translation and destabilizing mRNAs. They are produced by DICER-mediated processing of pre-miRNAs, which are short hairpin RNAs that are generated in the nucleus by the processing of primary RNA polymerase II transcripts (pri-miRNAs) by DROSHA (also known as RNase III); these pre-miRNAs are transported to the cytoplasm by exportin 5 (XPO5). Duplex miRNAs associate with Argonaute proteins (AGO1–AGO4), with subsequent loss of one RNA strand (the ‘passenger’ strand) to form miRNA-induced silencing complexes (mRISCs) in which the miRNA ‘guide’ strand is positioned with its seed sequence (comprising nucleotides 2–8) facing outwards, poised for interactions with targeted mRNAs. miR‑122 is one of ~1,900 recognized human miRNAs. It is among the most abundant miRNAs in hepatocytes and is conserved from zebrafish to primates84. miR‑122 is derived from pri-miR‑122, a 7.5 kb non-coding RNA transcribed from chromosome 18 under control of the hepatocyte-specific transcription factor HNF4α. The pri-miRNA is processed by DROSHA to an ~70 nucleotide pre-miR‑122 hairpin mir-122 gene (see the figure). Two copies of the miR‑122 guide strand bind to the 5ʹ end of the hepatitis C virus (HCV) positive-strand RNA ((+)RNA) genome in association with AGO2 (REFS 15,88). miR‑122 nucleotides Transcription 2–8 (the seed sequence; in bold in the figure) base pair with conserved tracts (S1 and S2) within the HCV genome sequence, whereas nucleotides 13–16 form additional accessory base pairs (see Pri-miR-122 the figure)91. Poly(rC)-binding protein 2 (PCBP2) binds to a sequence DROSHA overlapping the S2 site and competes with miR‑122 for binding to AAAAA the HCV 5ʹ untranslated region98. m7G Cropping 5ʹ C

C CUUAGCAG-AGCUGUGGAGUGUGACAAUGGUGUUUG- UGU U GGAUCGUCAUCGAUAAAUCACACUAUUACCGCAAAC CA A UAU A 3ʹ C

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A U G G G 3ʹ U 5ʹ ACAGU GUUUGU G GGUA GUGAGG G C GACACUCCACCAUGAAUCACUCCCCUG

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Polysome A large complex formed by multiple mature 80S ribosomes bound to an mRNA that is actively undergoing translation.

hypothesis, polysome analyses reveal a small but significant reduction in the fraction of HCV RNA associated with polysomes and engaged in translation following transfection of miR‑122 (REF. 98). Several possible mechanisms may explain how miR‑122 regulates the engagement of viral genomes in RNA synthesis versus translation. Electron microscopy and RNA pull-down experiments suggest that the HCV genome adopts a circularized structure through protein bridges forming between the 5ʹ and 3ʹ ends of the RNA in the presence of PCBP2 (REF. 72) or ILF3 (REF. 70), both of which are capable of binding the 5ʹ and 3ʹ ends of the RNA. As noted above, circularization of an RNA can enhance its translational efficiency by facilitating

AGO2 DICER Dicing and loading miR-122

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Nature Reviews | Microbiology

ribosome recycling (BOX 2). In addition, sequences within the HCV 3ʹ UTR can bind the 40S ribosome subunit independently of the IRES, providing a second possible mechanism by which circularization could enhance translation107. As miR‑122 competes with PCBP2 for binding to the extreme 5ʹ end of the genome at the S2 site74,98, miR‑122 might inhibit PCBP2‑mediated circularization of the genome and thereby reduce its engagement in translation (BOX 4). Consistent with this hypothesis, miR‑122 has no effect on viral RNA synthesis when transfected into cells previously depleted of PCBP2 (REF. 98). Alternatively, both SHAPE and atomic force microscopy suggest that the binding of miR‑122 induces changes in the tertiary RNA structure of the IRES95,108,



REVIEWS Box 4 | Does circularization of the hepatitis C virus genome regulate its engagement in translation versus replication? There is no conclusive proof that the hepatitis C virus (HCV) genome undergoes circularization, but various studies suggest PCBP2 bridge that this does occur and is mediated by poly(rC)-binding protein 2 (PCBP2), interleukin enhancer-binding factor 3 Ribosome (ILF3)70,72 and potentially other RNA-binding proteins that recycling interact with both the 5ʹ and 3ʹ untranslated regions (UTRs). (+)RNA Hypothetically, circularization could regulate the balance between RNA translation and synthesis. PCBP2 The figure displays a simplified model showing how the microRNA miR‑122 and PCBP2 could jointly regulate HCV genome circularization and thereby influence engagement of the RNA in the mutually incompatible processes of viral RNA synthesis and translation. Binding of the internal ribosome entry site (IRES)-transactivating factor PCBP2 to sequences Replicase 3ʹ near the 5ʹ and 3ʹ ends of the viral positive-strand RNA ((+)RNA) provides a protein bridge that promotes genome circularization72 and facilitates IRES-initiated translation, in 5ʹ Exosome part by enhancing the recycling of 40S subunits disengaging complex from the RNA after completing a round of translation (see the miR-122–AGO2 figure). The microRNA miR‑122 competes with PCBP2 for complex binding to the 5ʹ UTR98, promoting an open, noncircular genome conformation (see the figure). miR‑122 may also induce changes in the conformation of the IRES95. This leads to a reduction in translation and an increase in the initiation of negative-strand RNA ((–)RNA) synthesis, possibly owing to there now being a greater availability of the 3ʹ UTR for interactions with the HCV replicase98. miR‑122 acts in 3ʹ association with Argonaute 2 (AGO2) and also protects the 5ʹ 5ʹ end of the genome from 5ʹ‑to‑3ʹ exoribonuclease 1 (–) RNA (XRN1)‑mediated decay23. Interactions with multiple cellular proteins as well as the stable HCV RNA structure probably XRN1 protect the 3ʹ end from exosome-mediated decay when the genome is in the open conformation (see the figure). Long-range RNA–RNA interactions within the viral genome RNA synthesis may also be involved in the maintenance and/or induction of genome circularization (FIG. 1).

and this could also modulate the engagement of HCV RNA with the 40S ribosomal subunit. It should be noted that these two hypotheses are not mutually exclusive. In addition to PCBP2 and ILF3 (REFS 70,72), other host proteins that bind both ends of the viral genome — such as the LSM proteins LSM1 to LSM7, PTBP1, PTBP2, IGF2BP1 and SSB — could facilitate genome circularization68,69,77,109. Mobility shift and SHAPE studies also suggest the existence of direct RNA–RNA interactions that involve the 5ʹ and 3ʹ UTRs and may alter IRES conformation58,110 (FIG. 1). These RNA–RNA interactions are likely to be dynamic and, as in the case of EWSR1 binding to the CRE81, probably both influenced by and in turn influential to the interactions of the genome with host RNA-binding proteins. Such interactions could regulate the translational efficiency of the RNA and, thus, the balance of its engagement in protein synthesis versus RNA synthesis. RNA synthesis is also regulated by serine phosphorylation of the multifunctional NS5A protein. Several host kinases, including PI4KIIIα and casein kinase I isoform-α (CKIα), as well as multiple viral nonstructural proteins, including NS2 and NS4A, have been implicated in the generation of basal (56 kDa) and

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Nature Reviews | Microbiology

hyperphosphorylated NS5A species37,111. Mechanistic details are lacking, but hyperphosphorylation seems to have a negative effect on the capacity of NS5A to support replicase formation and RNA synthesis, while enhancing its movement to lipid droplets and promoting virion assembly 112,113. Hyperphosphorylation of NS5A is thus likely to deplete the intracellular (+)RNA pool. However, the impact of NS5A hyperphosphorylation on engagement of the residual RNA in translation versus RNA synthesis is unknown. Lipid peroxidation and the HCV replicase. A unique feature of the membranous web is its sensitivity to oxidative membrane damage14,114. HCV infection disturbs cellular redox homeostasis and induces a state of oxidative stress that is likely to be potentiated by inflammatory immune responses115. The presence of reactive oxygen species results in the peroxidation of polyunsaturated fatty acids within membranes. This lipid peroxidation causes both qualitative and quantitative changes in the membranous web14. A detailed molecular understanding of these processes is lacking, but it has been shown that peroxidation-resistant HCV variants selected in cell culture have mutations in membrane-proximal residues



REVIEWS of NS4A and NS5B14. Changes observed in the response to antivirals targeting the NS3‑4A protease and NS5B polymerase in peroxidation-sensitive virus, compared with peroxidation-resistant virus, suggest that the conformations of these proteins are altered by lipid peroxidation14, possibly through the formation of protein adducts, and that this results in impaired viral RNA synthesis. Lipid peroxidation limits the ability of HCV to replicate in cultured primary hepatocytes, an effect that can be reversed by lipid-soluble antioxidants such as vitamin E14. Notably, this phenotype is unique to HCV and is not observed in other (+)RNA viruses, including other members of the Flaviviridae family. The suppression of the HCV RNA synthetic capacity by oxidative membrane damage provides a novel autoregulatory mechanism that probably acts to maintain HCV replication at low levels, a feature that may favour viral evasion of immune surveillance and the survival of infected cells within the liver.

The yang of hepatitis C: RNA decay Eukaryotic cells are equipped with several distinct cytoplasmic RNA decay pathways that collectively act to maintain the proper abundance and integrity of mRNAs. These constitutively expressed decay pathways pose a barrier for the survival of RNA viruses and can be supplemented by ribonucleases that are induced as part of the cell-intrinsic innate immune response. In response to these challenges, RNA viruses, including HCV, have evolved a variety of strategies to evade mRNA decay programmes.

Flaviviridae A family of enveloped positive-strand RNA viruses comprising four genera: Flavivirus (dengue virus, yellow fever virus and others), Hepacivirus (hepatitis C virus), Pestivirus and Pegivirus.

P‑bodies (Processing bodies). Cytoplasmic aggregates of proteins engaged in the transport, storage and decay of host mRNA. These bodies represent foci of accumulation of 5ʹ‑to‑3ʹ exonuclease 1 (XRN1).

HCV RNA and host mRNA decay pathways. The degradation of host mRNAs takes place within the cytoplasm and begins with removal of the 3ʹ poly(A) tail by deadenylases, followed by exonucleolytic decay of the RNA body in either a 5ʹ-to-3ʹ or a 3ʹ-to-5ʹ direction116 (FIG. 3). In the 5ʹ-to-3ʹ decay pathway, the m7GpppN cap is removed by decapping enzymes117, and the mRNA body is then degraded by the cytoplasmic 5ʹ exoribonuclease, XRN1 (REF. 118). Alternatively, in 3ʹ-to-5ʹ decay, the mRNA body is degraded by the exosome complex, a multiprotein complex with both 3ʹ-to-5ʹ exonuclease and endonuclease activities119,120, and the residual cap structure is degraded by decapping scavenger enzyme (DCPS)121. Additional specialized mRNA decay machineries, including nonsense-mediated decay (NMD), no‑go decay and non-stop decay, provide surveillance over the integrity of cellular mRNAs116,122. Proteins involved in the NMD pathway restrict the replication of some (+)RNA viruses123, but none of these alternative RNA degradation pathways is known to contribute to HCV RNA decay. In addition to lacking a 5ʹ‑cap and a 3ʹ poly(A) tail, HCV RNAs are not synthesized in the nucleus and thus do not associate with transcription or splicing factors. These features distinguish HCV RNA from cellular mRNA and may thus target the viral RNA for degradation. However, the manner in which HCV RNA is degraded is dependent on its location in the cell and its functional status23. For example, when synthetic HCV RNA transcripts are electroporated into cells, they are degraded rapidly in both 5ʹ-to-3ʹ and 3ʹ-to-5ʹ directions

by XRN1 and the 3ʹ exosome complex, respectively, with a half-life (t1/2) of ~1.5–2.0 hours (REF. 23). By contrast, viral RNA is degraded much more slowly within infected cells after RNA synthesis is terminated with a specific inhibitor of the viral polymerase, in this case decaying with a t1/2 of 6–10 hours (REFS 23,124). This difference in the kinetics of RNA degradation is likely to reflect protection afforded to the RNA by the membranous web, or by packaging into nascent virions, in the infected cells. In these cells, HCV RNA is degraded primarily by the 5ʹ-to-3ʹ XRN1 pathway, with little or no contribution from the 3ʹ exosome complex 23. Consistent with this, only 5ʹ-to-3ʹ degradation intermediates have been identified within infected Huh‑7 cells23. Notably, XRN1 exclusively degrades RNAs possessing a 5ʹ monophosphate, and as HCV RNA terminates in a 5ʹ triphosphate, an as-yet‑unidentified cellular pyrophosphatase must contribute to XRN1‑mediated decay of the viral genome in infected cells23,125. Recent studies in infected cell cultures suggest that XRN1 is stalled by secondary RNA structure within the HCV 5ʹ UTR, and that this both represses XRN1 enzymatic activity and results in a global increase in host mRNA stability 126. However, similarly to the putative miR‑122 sponge effect described above93, it is not clear that such an effect occurs in vivo, where intracellular abundance of the viral genome is low. The primary role of XRN1 in the decay of HCV RNA is consistent with its involvement in the decay of other (+)RNA virus genomes127. However, a second 5ʹ exoribonuclease, XRN2, has also been implicated in the decay of HCV RNA128. XRN1 and XRN2 are very different proteins that share limited sequence homology only in their respective N‑terminal ribonuclease domains. XRN1 is expressed in the cytoplasm, where it is associated with P‑bodies (processing bodies) and functions in mRNA decay, whereas XRN2 is predominantly located in the nucleus, regulates termination of RNA polymerase II‑mediated transcription and contributes to the maturation of rRNA and snoRNA129,130. RNAi-mediated depletion of XRN2 can enhance HCV replication in cell culture, but this effect is limited to viruses that replicate robustly and generate cytopathic effects, such as the JFH1 isolate124,128. By contrast, XRN1 depletion enhances replication of multiple HCV strains, making XRN1 a more general restriction factor 23,124,131. XRN1 depletion also results in greater increases in the t1/2 of HCV RNA than are seen on XRN2 depletion, consistent with XRN1 having the primary role in decay 124. As HCV RNA does not localize to sites of XRN1 accumulation in P‑bodies, it is likely to be degraded by XRN1 within the cytosol23. Antiviral defence and HCV RNA decay. In addition to degradation via constitutive mRNA decay pathways, several IFN-inducible ribonucleases constitute alternative mechanisms for the degradation of HCV RNA132,133 (FIG. 3) . For example, the 2ʹ,5ʹ‑oligoisoadenylate synthetase-d ependent RNase L is a pivotal IFNinducible endoribonuclease that is important for intrinsic cellular defence against a variety of viruses. RNase L



REVIEWS a

m7G

mRNA AAAAAAAAAAAAAA 3ʹ

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AAA A 5ʹ-to-3ʹ decay

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Figure 3 | The yang of hepatitis C virus: RNA decay. a | Host mRNAs differ from Nature Reviews | Microbiology hepatitis C virus (HCV) RNA with respect to the presence of a 5ʹ 7‑methylguanylate cap 7 (m GpppN) and a 3ʹ poly(A) tail, both of which help to stabilize host cell mRNA. General mRNA decay initiates with removal of the 3ʹ poly(A) tail by a deadenylase complex and is followed by either 5ʹ‑to‑3ʹ or 3ʹ‑to‑5ʹ degradation. In the 3ʹ‑to‑5ʹ pathway, the RNA body is degraded by the 3ʹ exosome complex. The residual capped oligonucleotide is cleaved by decapping scavenger enzyme (DCPS). In the 5ʹ‑to‑3ʹ pathway, the 5ʹ‑cap is removed by decapping enzymes, and the RNA body is degraded by cytoplasmic 5ʹ‑to‑3ʹ exoribonuclease 1 (XRN1). b | Positive-strand HCV RNA possesses a 5ʹ triphosphate and terminates in a stable 3ʹ stem–loop. Its decay by constitutive mRNA decay pathways commences with removal of the 5ʹ triphosphate by an unknown cellular pyrophosphatase and proceeds via XRN1‑mediated degradation, as in the host mRNA 5ʹ‑to‑3ʹ decay pathway. However, HCV RNAs are protected from decay by their location within the detergent-resistant membranes of the membranous web and by interactions of the 5ʹ untranslated region with the microRNA miR‑122. Whether miR‑122 mediates protection against the putative pyrophosphatase or XRN1 (or both) is not known. Unlike decay of host mRNAs, the exosome complex has no role in this process. Interferon- and cytokine-inducible endoribonucleases and exoribonucleases represent alternative mechanisms for degradation of HCV RNA. Whereas these ribonucleases are actively expressed within the HCV-infected liver, their level of expression may be moderated by the ability of the HCV NS3‑4A protease to cleave key adaptor proteins involved in signalling from pathogen recognition receptors such as retinoic acid-inducible gene I (RIG‑I) and Toll-like receptors (TLRs), which are capable of sensing the presence of foreign RNA.

degrades single-strand RNAs of both viral and cellular origin, preferentially cleaving 3ʹ of UU and UA dinucleotides134. Although HCV induces little RNase L expression in infected cell cultures135, RNase L is abundant in liver tissue from patients with chronic HCV infection, and it colocalizes with NS5A, a component of the replicase136.

Purified RNase L cleaves HCV RNA in vitro137, and overexpression of RNase L in Huh‑7 cells suppresses HCV replication135. The lower-than-expected frequency of UA and UU dinucleotides in the HCV genome, together with the close proximity of such dinucleotides to stem– loop structures (providing an unfavourable context for RNase L cleavage), suggests that the HCV genome has evolved under selective pressure from RNase L53,137. Importantly, small RNA fragments generated by RNase L cleavage may induce signalling via RIG‑I that leads to activation of the IFNB promoter 136, thereby amplifying cell-intrinsic host defences. IFN-stimulated gene 20 kDa protein (ISG20) is a second IFN-stimulated ribonuclease133 that possesses 3ʹ exonuclease activity and restricts the replication of a broad range of RNA viruses138. Overexpression of ISG20 suppresses HCV replication in cell culture, and this antiviral action is dependent on the exonuclease activity of the protein138,139. In addition, zinc-finger CCCH domain-containing 12A (ZC3H12A; also known as MCPIP1) is an endoribonuclease that is induced by pro-inflammatory cytokines such as tumour necrosis factor (TNF), interleukin‑6 (IL‑6) and IL‑1β, when these cytokines are released in the context of viral infection140. ZC3H12A degrades specific cytokine mRNAs, but also degrades HCV RNA and, when overexpressed, suppresses HCV replication141. Zinc-finger CCCH-type antiviral protein 1 (ZC3HAV1; also known as ZAP) is induced by type I IFN, binds to a wide range of viral RNAs, and both represses their translation and promotes their decay by recruiting host mRNA decay factors142. However, the role of ZC3HAV1 during HCV infection has yet to be studied. In addition to IFN- and cytokine-induced ribonucleases, DROSHA — a nuclear type III endoribonuclease involved in miRNA biogenesis (BOX 3) — can act independently of IFNs to restrict the replication of RNA viruses143. In such cases, infection induces translocation of DROSHA to the cytoplasm, where it cleaves viral RNA. However, DROSHA is required for the biogenesis of miR‑122, an important pro-virus host factor, and does not restrict HCV replication80. Viral antagonism of RNA decay. Viruses counteract cellular RNA decay mechanisms at several levels. For example, replicating HCV genomes are sequestered within the membranous web, which protects the RNA from decay machinery 30. This strategy is common to many RNA viruses and probably explains the slower degradation of replicating HCV genomes versus the rapid decay observed with electroporated synthetic RNA23. Another evasion strategy adopted by some viruses involves active disruption of constitutive cellular RNA decay pathways, either by targeting specific decay factors for cleavage by viral proteases or by stimulating their degradation by the proteasome144. However, there is no evidence that HCV relies on this strategy to avoid RNA degradation. HCV may, however, restrict the effects of IFN-induced ribonucleases by actively constraining intrinsic antiviral cellular responses. Thus, although the IFN-regulated exonuclease ISG20 is transcriptionally induced within the



REVIEWS infected liver 145, this response may be limited by the HCV NS3‑4A protease, which ablates signalling pathways involved in the induction of IFN responses by targeting specific adaptor proteins, mitochondrial antiviral signalling protein (MAVS) and TRIF (also known as TICAM1), for cleavage1. Importantly, the interactions of HCV with the innate immune system are complex, and their impact in vivo is not well understood; for a more complete discussion on this topic, see REF. 1. HCV also exploits its interaction with miR‑122 to protect its genome from 5ʹ exonucleolytic degradation15,23,100. This contrasts with conventional miRNA action, which typically promotes mRNA degradation. Transfection of miR‑122 slows the decay of electroporated synthetic HCV RNA, and it also reduces the rate of decline in HCV RNA abundance that is seen when infected cells are treated with a viral polymerase inhibitor 15. miR‑122 does not increase the stability of transfected synthetic RNAs to which a 5ʹ‑cap has been added, indicating that it protects against 5ʹ decay 15,23. Consistent with the primary role of XRN1 in mediating viral RNA decay, as described above, miR‑122 also has no effect on the rate of HCV RNA decay in cells depleted of XRN1 (REF. 23). Furthermore, miR‑122‑mediated stabilization of the viral RNA requires transfection of duplex miR‑122 that is capable of being loaded into AGO2 and forming a RISC complex, and was not observed in Ago2–/– mouse embryonic fibroblasts15. Therefore, the viral RNA is stabilized by a miR‑122–AGO2 complex (BOX 3). Interestingly, miR‑122 stabilizes synthetic HCV RNA when incubated in HeLa cell lysates, in conditions in which HCV RNA degradation is mediated predominantly by the 3ʹ exosome complex 23. This remains unexplained mechanistically but suggests that the binding of miR‑122 influences long-range RNA–RNA interactions that have been proposed to exist between the 5ʹ and 3ʹ ends of the genome58,110, such that miR‑122 provides protection against 3ʹ exonucleolytic decay. Although the influence of miR‑122 on HCV RNA stability has been studied in detail, there could well be other factors that stabilize the viral genome. Two proteins known to bind HCV RNA are well-documented host mRNA stability factors. ELAVL1, which binds to U‑rich or AU‑rich elements in cellular mRNAs to enhance their stability, also binds the HCV 3ʹ UTR76. Similarly, PCBP2 stabilizes α‑globin mRNA in red blood cells146 and binds both the 5ʹ and 3ʹ UTRs of HCV RNA72. Depleting either PCBP2 or ELAVL1 impairs HCV replication72,76,80,147, but the influence of these proteins on HCV RNA stability has not been well studied.

estimated to release on average 25 virions per day, if all hepatocytes were to be infected. However, as sensitive assays detect viral RNA or proteins in only a minority (1–50%) of hepatocytes, the number of virions released by each infected hepatocyte is likely to be substantially higher 7–9. Fewer than 100 RNA copies are present in each infected hepatocyte (much fewer than in commonly used cell culture systems)8, probably representing only a few functional replication complexes in most cells9. Infected cells are not dispersed within the hepatic parenchyma but are grouped in clusters of variable size. RNA abundance is highest near the centre of the clusters, suggesting a dynamic in which cells in the periphery have been more recently infected7,150. However, a steady-state interpretation is also possible, such that replication in cells near the periphery of clusters is constrained by the expression of IFN-stimulated genes9. Modelling studies suggest that the t1/2 of infected cells is in the range of only 4–7 days (REF. 148), but this is based on several assumptions and a paucity of hard data. Nonetheless, even with the uncertainties in these estimates, it is evident that a balance must exist at the single-cell level between the production of new HCV genomes and their loss through a combination of decay and virion assembly and export (FIG. 2). It is uncertain what proportion of newly synthesized RNA genomes are exported from hepatocytes in nascent virions and what proportion decays within the cell, but the balance between RNA synthesis and decay is known to be determined through interactions of the viral genome with multiple host RNA-binding proteins and miR‑122, as discussed above. miR‑122 plays a key part in this process, promoting replication by optimizing the balance of genomes engaged in RNA synthesis versus translation98. miR‑122 also prevents the 5ʹ decay of (+)RNA15,23, adding to the protective effects of the membranous web and the potential viral blockade of IFNinducible ribonucleases. Host RNA-binding proteins (specifically, PCBP2 and ELAV1) could also contribute to the stability of the viral genome. Together, these actions favour viral replication and enhance the production and cellular abundance of viral RNA. However, this increase in viral abundance is balanced by the unique sensitivity of HCV to oxidative membrane damage, which limits the efficiency of the replicase complex and acts as a regulator of replication14. The net outcome of this conflict between synthesis and decay versus export of the viral genome is a very low abundance of viral proteins and RNAs within individual infected hepatocytes7,9, which favours cell survival and long-term viral persistence.

Yin and yang: balancing RNA synthesis and decay Long-term steady-state persistence of HCV requires a balance between the synthesis and decay of viral RNA at the level of the infected host. Mathematical modelling suggests that virions are cleared from the circulation with a t1/2 of ~45 minutes and that about 5 × 1012 virions are released from the liver each day 148. Much less is known about the dynamics of infection within single hepatocytes. As there are about 2 × 1011 hepatocytes in the adult human liver 149, each hepatocyte could be

Outlook Recent gains in understanding the functional impact of interactions between the HCV genome, host RNAbinding proteins and miR‑122 are beginning to provide insights into how HCV regulates the involvement of its genome in the competing processes of viral RNA translation and synthesis, and how this contributes to the overall balance of synthesis versus decay of the viral genome. miR‑122 is now known to directly stimulate viral RNA synthesis, resulting in secondary increases



REVIEWS in viral protein production98. miR‑122 also stabilizes the viral genome by preventing XRN1‑mediated 5ʹ RNA decay 15,23, thereby countering the restrictive effects of constitutive host RNA decay machinery. Notably, this action of miR‑122 is completely contrary to the canonical destabilizing action of miRNAs on host mRNAs. The absence of 3ʹ decay in the degradation of HCV RNA also distinguishes it from cellular mRNAs23. Thus, the yin and yang of HCV (+)RNA is unique. However, our understanding of both its synthesis and its decay is far from complete, and we should anticipate that other host factors contribute to these processes. Several important questions remain to be answered. For example, following the shut-off of new RNA synthesis, the rate of loss of viral RNAs is substantial and cannot be attributed entirely to XRN1, XRN2 or the exosome Horner, S. M. & Gale, M. Jr. Regulation of hepatic innate immunity by hepatitis C virus. Nat. Med. 19, 879–888 (2013). 2. Walker, C. M. Adaptive immunity to the hepatitis C virus. Adv. Virus Res. 78, 43–86 (2010). 3. Bartenschlager, R., Lohmann, V. & Penin, F. The molecular and structural basis of advanced antiviral therapy for hepatitis C virus infection. Nat. Rev. Microbiol. 11, 482–496 (2013). 4. Holmberg, S. D., Spradling, P. R., Moorman, A. C. & Denniston, M. M. Hepatitis C in the United States. N. Engl. J. Med. 368, 1859–1861 (2013). 5. Thomas, D. L. et al. The natural history of hepatitis C virus infection: host, viral, and environmental factors. JAMA 284, 450–456 (2000). 6. Prokunina-Olsson, L. et al. A variant upstream of IFNL3 (IL28B) creating a new interferon gene IFNL4 is associated with impaired clearance of hepatitis C virus. Nat. Genet. 45, 164–171 (2013). 7. Liang, Y. et al. Visualizing hepatitis C virus infections in human liver by two-photon microscopy. Gastroenterology 137, 1448–1458 (2009). 8. Kandathil, A. J. et al. Use of laser capture microdissection to map hepatitis C virus-positive hepatocytes in human liver. Gastroenterology 145, 1404–1413 (2013). 9. Wieland, S. et al. Simultaneous detection of hepatitis C virus and interferon stimulated gene expression in infected human liver. Hepatology 59, 2121–2130 (2014). 10. Teixeira, R., Marcos, L. A. & Friedman, S. L. Immunopathogenesis of hepatitis C virus infection and hepatic fibrosis: new insights into antifibrotic therapy in chronic hepatitis C. Hepatol. Res. 37, 579–595 (2007). 11. Westbrook, R. H. & Dusheiko, G. Natural history of hepatitis C. J. Hepatol. 61, S58–S68 (2014). 12. Walters, K. A. et al. Genomic analysis reveals a potential role for cell cycle perturbation in HCVmediated apoptosis of cultured hepatocytes. PLoS Pathog. 5, e1000269 (2009). 13. Kannan, R. P., Hensley, L. L., Evers, L., Lemon, S. M. & McGivern, D. R. Hepatitis C virus infection causes cell cycle arrest at the level of entry to mitosis. J. Virol. 85, 7989–8001 (2011). 14. Yamane, D. et al. Regulation of the hepatitis C virus RNA replicase by endogenous lipid peroxidation. Nat. Med. 20, 927–935 (2014). This paper describes the unique manner in which the HCV replicase is negatively regulated by cellular lipid peroxidation. 15. Shimakami, T. et al. Stabilization of hepatitis C virus RNA by an Ago2–miR‑122 complex. Proc. Natl Acad. Sci. USA 109, 941–946 (2012). 16. Zeisel, M. B., Felmlee, D. J. & Baumert, T. F. Hepatitis C virus entry. Curr. Top. Microbiol. Immunol. 369, 87–112 (2013). 17. Shulla, A. & Randall, G. Hepatitis C virus — host interactions, replication, and viral assembly. Curr. Opin. Virol. 2, 725–732 (2012). 18. Honda, M., Beard, M. R., Ping, L. H. & Lemon, S. M. A phylogenetically conserved stem-loop structure at the 5ʹ border of the internal ribosome entry site of hepatitis C virus is required for cap-independent viral translation. J. Virol. 73, 1165–1174 (1999). 1.

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Acknowledgements

The authors apologize to the many colleagues whose work they have been unable to cite and thank K. McKnight and A. Perelson for helpful discussions. This work was supported in part by grants from the US National Institutes of Health (R01‑AI095690 and R01‑CA164029) and the University of North Carolina Cancer Research Fund.

Competing interests statement

The authors declare competing interests: see Web version for details.


Prevalence of hepatitis C virus antibodies and hepatitis C virus-RNA in an urban population.

Detection of serum hepatitis C virus RNA.

Hepatitis C virus RNA in saliva.

Hepatitis C virus RNA and hepatitis C virus antibody in the serum of patients with abnormal liver function.

Hepatitis C virus RNA and antibody response in the clinical course of acute hepatitis C virus infection.

[The hepatitis C virus].

Correlation between the serum level of hepatitis C virus RNA and disease activities in acute and chronic hepatitis C.

[Analysis of hepatitis C virus RNA from semen and urine of the patients with chronic type C hepatitis].

The role of microRNAs in hepatitis C virus RNA replication.

Total synthesis and anti-hepatitis C virus activity of MA026.

Use of aminotransferase, hepatitis C antibody, and hepatitis C polymerase chain reaction RNA assays to establish the diagnosis of hepatitis C virus infection in a diagnostic virology laboratory.

Hepatitis C virus and autophagy.

Hepatitis C virus and autoimmunity.

Cryoglobulinaemia and hepatitis C virus.

Hepatitis C virus and lymphoma.

Hepatitis B virus and hepatitis C virus dual infection.

Hepatitis C virus.

Hepatitis C virus.

Association of hepatitis C virus carrier state with the occurrence of hepatitis C virus core antibodies.

Hepatitis C virus replication.

Hepatitis C virus upstanding.

Detection of hepatitis C virus RNA by in situ hybridization.

[Hepatitis C virus].

Hepatitis C virus RNA detection in serum and peripheral blood mononuclear cells of patients with hepatitis C.